Funded by the NSF (0854551).

1. Pre-experimental Preparations
<ol>
	<li>Prepare six solutions (see Table 1 for compositions): (a) 100 ml NZR (normal zebrafish Ringer's), (b) 50 ml NZR + Tricaine (MS222), (c) 10 ml NZR +BSA; (d) 100 ml LoCaS (low Ca2+ solution), (e) 10 ml LoCaS + papain, (f) 100 ml K+-internal solution. Solutions (a), (b), (d) and (f) can be stored for up to one month at 4 &deg;C. Solutions (c) and (e) should be prepared the day of experimentation. All solutions should be at room temperature before beginning experiments.</li>
	<li>Label and fill four 35 mm Petri dishes approximately halfway with solutions (a), (c), (d) and (e).</li>
	<li>Make a dissecting dish by filling a 60 mm Petri dish with Sylgard (Dow Corning, Midland, MI) and then cutting a cavity into it that will allow a fish to sit upright without tipping over.</li>
	<li>Prepare at least two cell isolation tools by gluing the follicle end of a dog hair to the end of a glass Pasteur pipette with superglue, allowing the hair to extend past the end of the pipette by approximately 0.5 cm. The hairs from soft-furred dogs like Labrador Retrievers and Weimaraners work well.</li>
</ol>
2. Isolation of Auditory and Vestibular Labyrinth
<ol>
	<li>Sacrifice one adult zebrafish by immersion in a beaker containing NZR + Tricaine. Visually monitor the gills until all opercular movements have ceased. Wait an additional ten minutes before removing fish and rinsing with fresh NZR. These procedures have been approved by the Institutional Animal Care and Use Committee (IACUC) of Pepperdine University but approval should be obtained from your own institution.</li>
	<li>Pin the fish to the dissecting dish dorsal side up by inserting one standard sewing straight pin (approximately 2.5 cm long) through each eye socket and a third pin through the dorsal-ventral axis about one-third to one-half of the distance from head to tail as illustrated in Figure 1A.</li>
	<li>Under a zoom (0.62-5x) stereo dissecting microscope fitted with 10x eyepieces, use spring scissors (Fine Science Tools catalogue number 15000-02) to expose the brain and a short segment of the spinal cord by removing the skull roof from a point approximately 0.5 cm caudal to the level of the gills forward to the nose of the fish. Cut the spinal cord and lift the brain with fine forceps (Fine Science Tools catalogue number 11251-30) while cutting the spinal nerves and other attachments; remove the brain and discard (Figure 1B).</li>
	<li>The ventral half of the skull capsule that remains forms a "bowl" that contains the inner ear organs. Observe the prominent, white, opaque otoliths in the lagenas and sacculi located symmetrically at the caudal end of each inner ear and the laterally positioned utricle otoliths more rostrally (Figures 1C and 4A).</li>
	<li>Remove the bowl of bone by cutting all ventral and lateral attachments while gently lifting it out of the head with fine forceps. Place it into the Petri dish containing LoCaS (Figures 1D and 4B).</li>
	<li>Using two pairs of fine forceps, crack apart the bowl at its midline by either grabbing the lateral edges of the bone and pulling them apart or alternatively, separating the right and left halves by inserting the points of the forceps in the center and prying the halves apart. Remove the two labyrinths from the bone using fine forceps (Figure 2).</li>
	<li>Inspect the labyrinths to identify the auditory and vestibular end-organs: the semicircular canals, utricles, sacculi and lagenas (Figure 4C). The pinkish areas under each otolith are the maculae containing the hair cells. Similarly, a strip of pink inside the ampullae of the semicircular canals identifies the cupula where the hair cells are located.</li>
	<li>Carefully remove the otoliths from the lagena and utricle using the fine forceps (Figures 2 and 4C). It is not necessary to remove the otolith from the saccule as doing so may damage the saccular hair cells.</li>
</ol>
3. Hair Cell Isolation
<ol>
	<li>Using a plastic transfer pipette (Fisher Scientific catalogue number 13-711-9AM), move the end organs to the dish containing LoCaS + papain minimizing the volume of fluid transferred.</li>
	<li>Incubate these structures in this solution for thirty minutes at room temperature.</li>
	<li>At the end of the incubation period, move the structures to the dish containing NZR + BSA and incubate in this solution for at least thirty minutes. Hair cells will remain healthy in this solution for up to four hours but will deteriorate after 1-2 hours after isolation (see next steps).</li>
	<li>In preparation for cell isolation, move one of the end-organs to the dish containing NZR.</li>
	<li>For the lagenas and utricles, use a dog hair to gently lift off the maculae-sheets of pinkish epithelial tissue that contain the hair cells (Figure 3A). The maculae can be scooped out (using a dog hair) from under the otoliths of the sacculi, and the cupulae from their locations inside the ampullae of the semicircular canals.</li>
	<li>Using two cell isolation tools prepared in step 1.4, triturate a macula or cupula to generate a debris field that contains the hair cells (Figure 3B). Allow the cells at least five minutes to settle onto the bottom of the dish before moving it.</li>
	<li>Relocate the dish to the stage of an inverted microscope equipped with phase contrast optics. Observe cells under 40X magnification and note the diversity in morphology. Many cells are avocado-shaped while others are long and thin. The presence of apical hair bundles identifies them as hair cells and phase-brightness assures their health. Photomicrographs of isolated hair cells are shown in Figure 5.</li>
</ol>
4. Patch Clamping Zebrafish Hair Cells
<ol>
	<li>In preparation for perforated patch recordings5,6, prepare amphotericin-containing internal solution as follows: place 5 mg amphotericin B (Sigma A4888) into a 1.5 ml microcentrifuge tube and fill with 100 &mu;l DMSO. Immediately vortex this solution for 10-20 seconds until all of the amphotericin is dissolved. Then, pipette 625 &mu;l K+-internal solution into a second microcentrifuge tube. Add 10 &mu;l of the amphotericin solution to the K+-internal solution and vortex as above. The stock amphotericin solution will last several hours but discard the final solution after one hour.</li>
	<li>Fabricate patch pipettes from BF 150-86-10 borosilicate glass (Sutter Instruments) using a multi-stage puller (Sutter P-1000). Pipettes should have tip diameters of approximately 2 &mu;m and will have resistances of 1-3 M&Omega; with the solutions in Table 1. The settings on the puller that work well are as follows: Heat=Ramp minus10, Pull=0, Velocity=18, Delay=1, Pressure=600. The shape of the tip of a good pipette is evident from the photograph in the inset c of Figure 5. Blunt "bullet-shaped" pipettes are preferred as they offer the least access resistance during electrophysiological recordings.</li>
	<li>Use a 28 gauge Microfil syringe needle (MF28G-5; World Precision Instruments) to fill a recording electrode halfway with the amphotericin-containing K+-internal solution.</li>
	<li>Affix the electrode to the headstage of a patch clamp amplifier. Apply -10 mV, 10 ms voltage steps at 10 Hz and maintain positive pressure in the electrode as it is maneuvered through the air/solution interface and down to the bottom of the recording dish near the cells.</li>
	<li>Position the electrode orthogonally to a healthy looking cell. When the electrode is near enough to the cell so that it begins move away from the out-flowing solution, quickly reverse the pressure, applying a slight vacuum to the electrode until the cell "jumps" onto it. Immediately cease suction on the electrode and apply a holding potential of -70 mV to the inside of the electrode. Within a few seconds a gigaohm seal will form.</li>
	<li>Observe the capacitive current transients at the onset and termination of the voltage steps that appear soon after seal formation. Continue to monitor the perforation of the membrane under the electrode by the amphotericin by watching as the transients grow in magnitude to their maximum size (in about ten minutes).</li>
	<li>To confirm the establishment of whole-cell (perforated) configuration in a healthy cell, elicit outward K+ currents by applying hyperpolarizing and depolarizing steps of voltage. Most hair cells have prominent outward currents (see Figure 6).</li>
</ol>
5. Representative Results
Figure 4 shows images taken during the steps of cell isolation. In panel A, the otoliths associated with the lagenas, saccluli and utricles can be seen through the thin layer of bone that overlies them. These opaque structures provide convenient landmarks to aid in the safe removal of the end organs from the animal during the subsequent steps of the dissection. Panel B shows the "bowl" of bone that contains the intact labyrinths and the prominent otoliths in the utricles, lagenas and sacculi. Panel C shows a labyrinth that has been removed from the bone. Note the large otolith with the scalloped edge in the lagena, the icicle-shaped otolith in the saccule and the bean-shaped otolith in the utricle. Some of the diversity in size and morphology seen in cells isolated from the lagena is illustrated in Figure 5. A healthy cell is readily identified as being phase bright with a sharp perimeter. Cell shapes can be roughly divided into two classes: avocado-shaped (avo) and long and thin (thn) although the sizes of cells within each group can vary markedly (compare for example avo 1 and avo 3). Inset a shows a cluster of three cells that were not completely isolated from one another. The black color and granular appearance of the cell in inset b readily identifies this cell as dead. Inset c shows the tip of a patch pipette illustrating the shape and dimensions appropriate for recording from these cells.
The current traces shown in Figure 6 were obtained in response to hyperpolarizing and depolarizing steps of potential imposed on a lagena cell similar to the avo 2 shown in Figure 5. Current responses in cells of different sizes and shapes can vary widely suggesting diversity in the complement of voltage-gated channels.
Figures 1-3: Cartoons illustrating steps in the isolation of hair cells.
<img alt="Figure 1" src="/files/ftp_upload/4281/4281fig1.jpg" /> 
 Figure 1. Removal of the skull capsule containing the inner ear labyrinths from zebrafish. A. Pin sacrificed zebrafish dorsal side up to dissecting dish. B. Open skull, remove and discard brain. C. Observe otoliths in utricles, lagenas and sacculi. D. Remove ventral portion of skull capsule containing the labyrinths.
<img alt="Figure 2" src="/files/ftp_upload/4281/4281fig2.jpg" /> 
 Figure 2. Removal of labyrinths from skull capsule. A. Crack apart skull capsule at its midline. B. Remove labyrinths from bone. C. Remove the otoliths from the lagenas and utricles using forceps.
<img alt="Figure 3" src="/files/ftp_upload/4281/4281fig3.jpg" /> 
 Figure 3. Isolation of hair cells from labyrinths. A. Gently lift off maculae with a dog hair. B. Triturate maculae with two dog hairs. C. Use isolated hair cells for patch clamp.
<img alt="Figure 4" src="/files/ftp_upload/4281/4281fig4.jpg" /> 
 Figure 4. Images taken during steps in the isolation of individual cells. A. After removal of the brain, the otoliths associated with the two lagenas and sacculi (arrows) and utricles (arrowheads) can be visualized. B. After removal of the ventral portion of the skull capsule that contains the inner ear organs from the animal. The cartoon in the right half of B identifies the locations of the left utricle, lagena and saccule. The dashed line indicates the approximate position of the left labyrinth. C. Right labyrinth (medial view) D: Cartoon drawing illustrating key portions of labyrinth in C. a: anterior semicircular canal, b: horizontal canal, c: posterior canal, d: utricular otolith, e: saccular otolith, f: lagenal otolith. Scale bar in D represents 1 mm for A and B, 0.5 mm for C and D.
<img alt="Figure 5" src="/files/ftp_upload/4281/4281fig5.jpg" /> 
 Figure 5. Isolated, healthy cells from the lagena illustrating the diversity of morphology; avo: avocado shaped cells; thn: long, thin cells. Inset a: cluster of three incompletely isolated cells; inset b: dead cell; inset c: tip of patch electrode used in electrophysiological recordings from zebrafish hair cells. Scale bar = 20 &mu;m applies to the main figure and all insets.
<img alt="Figure 6" src="/files/ftp_upload/4281/4281fig6.jpg" /> 
 Figure 6. Currents recorded in a patch clamped hair cell. Averaged responses in a cell (similar to avo 2 in Figure 5) to three presentations of voltage steps applied in 10 mV increments from -140 mV to +70 mV from a holding potential of -70 mV. Note the inactivating current that appears at more depolarized potentials. Voltage step magnitudes are shown next to some of the traces.

<ol>
	<li>Lewis, R. S., Hudspeth, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Voltage-+and+ion-dependent+conductances+in+solitary+vertebrate+hair+cells.">Voltage- and ion-dependent conductances in solitary vertebrate hair cells.</a> Nature. 304, 538-5341 (1983).</li><li>Art, J. J., Fettiplace, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Variation+of+membrane+properties+in+hair+cells+isolated+from+the+turtle+cochlea.">Variation of membrane properties in hair cells isolated from the turtle cochlea.</a> J. Physiol. 385, 207-242 (1987).</li><li>Sugihara, I., Furukawa, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Morphological+and+functional+aspects+of+two+different+types+of+hair+cells+in+the+goldfish+sacculus.">Morphological and functional aspects of two different types of hair cells in the goldfish sacculus.</a> J. Neurophysiol. 62, 1330-1343 (1989).</li><li>Lee, S., Briklin, O., Hiel, H., Fuchs, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium-dependent+inactivation+of+calcium+channels+in+cochlear+hair+cells+of+the+chicken.">Calcium-dependent inactivation of calcium channels in cochlear hair cells of the chicken.</a> J. Physiol. 583, 909-922 (2007).</li><li>Horn, R., Marty, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Muscarinic+activation+of+ionic+currents+measured+by+a+new+whole-cell+recording+method.">Muscarinic activation of ionic currents measured by a new whole-cell recording method.</a> J. Gen. Physiol. 92, 145-159 (1988).</li><li>Rae, J. R., Cooper, K., Gates, P., Watsky, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Low+access+perforated+patch+recordings+using+amphotericin+B.">Low access perforated patch recordings using amphotericin B.</a> J. Neurosci. Meth. 37, 15-26 (1991).</li><li>Lumpkin, E. A., Hudspeth, A. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Detection+of+Ca2++entry+through+mechanosensitive+channels+localizes+the+site+of+mechanoelectrical+transduction+in+hair+cells.">Detection of Ca2+ entry through mechanosensitive channels localizes the site of mechanoelectrical transduction in hair cells.</a> Proc. Natl Acad. Sci. U.S.A. 92, 10297-10301 (1995).</li><li>Liang, J., Burgess, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Gross+and+fine+dissection+of+inner+ear+sensory+epithelia+in+adult+zebrafish+(Danio+rerio).">Gross and fine dissection of inner ear sensory epithelia in adult zebrafish (Danio rerio).</a> J. Vis Exp. 27, e1211 (2009).</li><li>Hudspeth, A. J., Lewis, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Kinetic+analysis+of+voltage-+and+ion-dependent+conductances+in+saccular+hair+cells+of+the+bull-frog,+Rana+catesbeiana.">Kinetic analysis of voltage- and ion-dependent conductances in saccular hair cells of the bull-frog, Rana catesbeiana.</a> J. Physiol. 400, 237-274 (1988).</li><li>Hudspeth, A. J., Lewis, R. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+model+for+electrical+resonance+and+frequency+tuning+in+saccular+hair+cells+of+the+bull-frog,+Rana+catesbeiana.">A model for electrical resonance and frequency tuning in saccular hair cells of the bull-frog, Rana catesbeiana.</a> J. Physiol. 400, 275-297 (1988).</li><li>Parsons, T. D., Lenzi, D., Almer, W., Roberts, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Calcium-triggered+exocytosis+in+an+isolated+presynaptic+cell:+capacitance+measurements+in+saccular+hair+cells.">Calcium-triggered exocytosis in an isolated presynaptic cell: capacitance measurements in saccular hair cells.</a> Neuron. 13, 875-883 (1994).</li><li>Kim, M. -. H., Vickers, E., von Gersdorff, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Patch-clamp+capacitance+measurements+and+Ca2++imaging+at+single+nerve+terminals+in+retinal+slices.">Patch-clamp capacitance measurements and Ca2+ imaging at single nerve terminals in retinal slices.</a> J. Vis Exp. 59, e3345 (2012).</li><li>Housley, G. D., Ashmore, J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Direct+measurement+of+the+action+of+acetylcholine+on+isolated+outer+hair+cells+of+the+guinea+pig+cochlea.">Direct measurement of the action of acetylcholine on isolated outer hair cells of the guinea pig cochlea.</a> Proc. R. Soc. Lond. B. 244, 161-167 (1991).</li><li>Nicolson, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+genetics+of+hearing+and+balance+in+zebrafish.">The genetics of hearing and balance in zebrafish.</a> Ann. Rev. Genetics. 39, 9-22 (2005).</li></ol>

No conflicts of interest declared.

Careful dissection of the end organs and gentle treatment of the cells is critical to the successful isolation of healthy, intact and physiologically active hair cells. Adhering to the following tips will assure success:
<ol>
 <li>It is easier to obtain healthy cells from smaller fish (2-2.5 cm in length). Larger fish have many more fat droplets that obscure visualization during the dissection process.</li>
 <li>Care must be taken when removing the otoliths, which overlie the hair cells. Avoid touching the cells when grabbing the otoliths with forceps. </li>
 <li>Best results are obtained when the maculae and cupulae are lifted off as sheets and the cells triturated from the epithelium.</li>
 <li>We find the use of a dog hair to triturate cells to be superior to other methods, including using human eyelashes7 which are not as tapered and less flexible.</li>
 <li>During patch clamping, application of positive pressure to the electrode as it passes through the air/solution interface is important to keep the tip clean. The release of the positive pressure to enable the cell to "jump" onto the electrode is also important. This method clears a small volume of amphotericin-containing solution away from the tip, thus increasing the likelihood of forming a gigaohm seal with the cell. In our experiments we use the perforated patch recording technique instead of the more traditional whole-cell configuration because of the fragility of the cells. Attempting to "go whole cell" often results in the loss of the gigaohm seal.</li>
</ol>
The method described here will yield hundreds of healthy hair cells per animal. The technique can be performed at room temperature and requires no special equipment beyond a dissecting microscope. A previous report in JoVE describes the dissection of the inner ear in zebrafish using a ventral approach8. These authors demonstrate the dissection of whole, paraformaldehyde-fixed inner ear organs. We encourage the reader to watch this video for comparison with our methods. One advantage of the methods presented here is the obtaining of live cells useful for physiological investigations. Besides their use in patch clamp experiments to study the voltage-gated channels (as shown here) the use of these cells can be extended to study cell resonance9,10, monitor neurotransmitter release by making capacitance measurements11,12, investigate neuromodulation induced by the activation of ligand gated channels13 as well as tap into the wealth of information that can be gleaned from using mutants that affect both hearing and balance14.

<table border="1">
 <tbody>
 <tr>
 <td>Solution Name</td>
 <td>Recipe</td>
 </tr>
 <tr>
 <td>(a) NZR (Normal Zebrafish Ringer's) (in mM)</td>
 <td>116 NaCl, 2 KCl, 2 CaCl2, 3 Glucose, 5 Na+-HEPES, pH 7.35.</td>
 </tr>
 <tr>
 <td>(b) NZR + Tricaine</td>
 <td>NZR + 0.02% 3-aminobenzoic acid ethyl ester methane sulfonate (Sigma A5040).</td>
 </tr>
 <tr>
 <td>(c) NZR + BSA</td>
 <td>NZR + 0.4% Bovine Serum Albumin (Sigma A2153). </td>
 </tr>
 <tr>
 <td>(d) LoCaS (Low Ca2+ Solution) (in mM)</td>
 <td>100 NaCl, 2 KCl, 0.05 CaCl2, 0.05 MgCl2, 3 Glucose, 30 Na+-HEPES, pH 7.35. </td>
 </tr>
 <tr>
 <td>(e) LoCaS + papain</td>
 <td>LoCaS + 0.05% L-cysteine (Sigma C1276) + 0.2% papain (Sigma P3375).</td>
 </tr>
 <tr>
 <td>(f) K+-internal solution (in mM)</td>
 <td>52 K2SO4, 38 KCl, 1 K+-EGTA, 5 K+-HEPES, pH 7.3.</td>
 </tr>
 </tbody>
</table>
Table 1. Solutions.

patch clamp recordings in inner ear hair cells isolated from zebrafish

Patch clamp analyses of the voltage-gated channels in sensory hair cells isolated from a variety of species have been described previously1-4 but this video represents the first application of those techniques to hair cells from zebrafish. Here we demonstrate a method to isolate healthy, intact hair cells from all of the inner ear end-organs: saccule, lagena, utricle and semicircular canals. Further, we demonstrate the diversity in hair cell size and morphology and give an example of the kinds of patch clamp recordings that can be obtained. The advantage of the use of this zebrafish model system over others stems from the availability of zebrafish mutants that affect both hearing and balance. In combination with the use of transgenic lines and other techniques that utilize genetic analysis and manipulation, the cell isolation and electrophysiological methods introduced here should facilitate greater insight into the roles hair cells play in mediating these sensory modalities.

Watch this Scientific Journal Video about Patch Clamp Recordings in Inner Ear Hair Cells Isolated from Zebrafish at JoVE.com

Patch Clamp Recordings in Inner Ear Hair Cells Isolated from Zebrafish

The purpose of this video is to demonstrate procedures for obtaining healthy, intact hair cells from the inner ear organs of adult zebrafish and then using them for patch clamp studies aimed at characterizing the biophysical properties of their voltage-gated channels.

Patch  clamp analyses of the voltage-gated channels in sensory hair cells isolated  from a variety of species have been described previously1-4 but this  ...

patch-clamp-recordings-in-inner-ear-hair-cells-isolated-from-zebrafish

Research

JoVE Journal

Neuroscience

16.6K Views.  Pepperdine University. The purpose of this video is to demonstrate procedures for obtaining healthy, intact hair cells from the inner ear organs of adult zebrafish and then using them for patch clamp studies aimed at characterizing the biophysical properties of their voltage-gated channels.

Video: Patch Clamp Recordings in Inner Ear Hair Cells Isolated from Zebrafish

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Paired Patch Clamp Recordings from Motor-neuron and Target Skeletal Muscle in Zebrafish

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle. This is a direct consequence of the accessibility to both cell types and ability to visually distinguish the single segmental CaP motor-neuron on the basis of morphology and location. This video demonstrates the microscopic methods used to identify a CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type. Identification of the CaP motor-neuron type is confirmed by either dye filling or by the biophysical features such as action potential waveform and cell input resistance. Motor-neuron recordings routinely last for one hour permitting long-term recordings from multiple different target muscle cells. Control over the motor-neuron firing pattern enables measurements of the frequency-dependence of synaptic transmission at the neuromuscular junction. Owing to a large quantal size and the low noise provided by whole cell voltage clamp, all of the unitary events can be resolved in muscle. This feature permits study of basic synaptic properties such as release properties, vesicle recycling, as well as synaptic depression and facilitation. The advantages offered by this in vivo preparation eclipse previous neuromuscular model systems studied wherein the motor-neurons are usually stimulated by extracellular electrodes and the muscles are too large for whole cell patch clamp. The zebrafish preparation is amenable to combining electrophysiological analysis with a wide range of approaches including transgenic lines, morpholino knockdown, pharmacological intervention and in vivo imaging. These approaches, coupled with the growing number of neuromuscular disease models provided by mutant lines of zebrafish, open the door for new understanding of human neuromuscular disorders.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target skeletal muscle. This video demonstrates the microscopic methods used to identify a segmental CaP motor-neuron and target muscle cells as well as the methodologies for recording from each cell type.

Larval zebrafish represent the first vertebrate model system to allow simultaneous patch clamp recording from a spinal motor-neuron and target muscle.  ...

Postsynaptic Recordings at Afferent Dendrites Contacting Cochlear Inner Hair Cells: Monitoring Multivesicular Release at a Ribbon Synapse

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the postsynaptic activity of a single ribbon synapse 1-4. Ribbon synapses of sensory cells release neurotransmitter continuously, the rate of which is modulated in response to graded changes in IHC membrane potential 5. Ribbon synapses have been shown to operate by multivesicular release, where multiple vesicles can be released simultaneously to evoke excitatory postsynaptic currents (EPSCs) of varying amplitudes 1, 4, 6-11. Neither the role of the presynaptic ribbon, nor the mechanism underlying multivesicular release is currently well understood.
The IHC is innervated by 10-20 auditory nerve fibers, and every fiber contacts the IHC with a unmyelinated single ending to form a single ribbon synapse. The small size of the afferent boutons contacting IHCs (approximately 1 &mu;m in diameter) enables recordings with exceptional temporal resolution to be made. Furthermore, the technique can be adapted to record from both pre- and postsynaptic cells simultaneously, allowing the transfer function at the synapse to be studied directly 2. This method therefore provides a means by which fundamental aspects of neurotransmission can be studied, from multivesicular release to the elusive function of the ribbon in sensory cells.

Whole-cell patch-clamp recordings from auditory nerve fiber dendrites at the inner hair cell ribbon synapse in the mammalian cochlea.

The afferent synapse between the inner hair cell (IHC) and the auditory nerve fiber provides an electrophysiologically accessible site for recording the ...

Isolating LacZ-expressing Cells from Mouse Inner Ear Tissues using Flow Cytometry

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues presents a unique challenge because of the paucity of cells of interest and few transgenic reporter mouse models. Here, we describe a protocol using fluorescence-conjugated probes to selectively label LacZ-positive cells from the neonatal cochleae.
The most common underlying pathology of sensorineural hearing loss is the irreversible damage and loss of cochlear sensory hair cells, which are required to transduce sound waves to neural impulses. Recent evidence suggests that the murine auditory and vestibular organs harbor stem/progenitor cells that may have regenerative potential1,2. These findings warrant further investigation, including identifying specific cell types with stem/progenitor cell characteristics. The Wnt signaling pathway has been demonstrated to play a critical role in maintaining stem/progenitor cell populations in several organ systems3-7. We have recently identified Wnt-responsive Axin2-expressing cells in the neonatal cochlea, but their function is largely unknown8.
To better understand the behavior of these Wnt-responsive cells in vitro, we have developed a method of isolating Axin2-expressing cells from cochleae of Axin2-LacZ reporter mice9. Using flow cytometry to isolate Axin2-LacZ positive cells from the neonatal cochleae, we could in turn execute a variety of experiments on live cells to interrogate their behavior as stem/progenitor cells. Here, we describe in detail the steps for the microdissection of neonatal cochlea, dissociation of these tissues, labeling of the LacZ-positive cells using a fluorogenic substrate, and cell sorting. Techniques for dissociating cochleae into single cells and isolating cochlear cells via flow cytometry have been described2,10-12. We have made modifications to these techniques to establish a novel protocol to isolate LacZ-expressing cells from the neonatal cochlea.

Flow cytometry is a powerful tool allowing for the isolation and study of specific cell populations. This protocol describes steps for isolating LacZ-expressing cells from cochlear tissues from neonatal transgenic mice. Dissociated cochlear cells were labeled using fluorescent-conjugated substrates of &#946;-galactosidase prior to separation via flow cytometry.

Isolation of specific cell types allows one to analyze rare cell populations such as stem/progenitor cells. Such an approach to studying inner ear tissues ...

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and the organ of Corti in the coiled cochlea. The cristae and maculae contain vestibular hair cells that transduce mechanical stimuli to subserve the special sense of balance, while auditory hair cells in the organ of Corti are the primary transducers for hearing 1. Cell fate specification in these sensory epithelia and morphogenesis of the semicircular canals and cochlea take place during the second week of gestation in the mouse and are largely completed before birth 2,3. Developmental studies of the mouse inner ear are routinely conducted by harvesting transgenic embryos at different embryonic or postnatal stages to gain insight into the molecular basis of cellular and/or morphological phenotypes 4,5. We hypothesize that gene transfer to the developing mouse inner ear in utero in the context of gain- and loss-of-function studies represents a complimentary approach to traditional mouse transgenesis for the interrogation of the genetic mechanisms underlying mammalian inner ear development6.
The experimental paradigm to conduct gene misexpression studies in the developing mouse inner ear demonstrated here resolves into three general steps: 1) ventral laparotomy; 2) transuterine microinjection; and 3) in vivo electroporation. Ventral laparotomy is a mouse survival surgical technique that permits externalization of the uterus to gain experimental access to the implanted embryos7. Transuterine microinjection is the use of beveled, glass capillary micropipettes to introduce expression plasmid into the lumen of the otic vesicle or otocyst. In vivo electroporation is the application of square wave, direct current pulses to drive expression plasmid into progenitor cells8-10. 
We previously described this electroporation-based gene transfer technique and included detailed notes on each step of the protocol11. Mouse experimental embryological techniques can be difficult to learn from prose and still images alone. In the present work, we demonstrate the 3 steps in the gene transfer procedure. Most critically, we deploy digital video microscopy to show precisely how to: 1) identify embryo orientation in utero; 2) reorient embryos for targeting injections to the otocyst; 3) microinject DNA mixed with tracer dye solution into the otocyst at embryonic days 11.5 and 12.5; 4) electroporate the injected otocyst; and 5) label electroporated embryos for postnatal selection at birth. We provide representative examples of successfully transfected inner ears; a pictorial guide to the most common causes of otocyst mistargeting; discuss how to avoid common methodological errors; and present guidelines for writing an in utero gene transfer animal care protocol.

Gene Transfer to the Developing Mouse Inner Ear by In Vivo Electroporation

The mouse inner ear is a placode-derived sensory organ whose developmental program is elaborated during gestation. We define an in utero gene transfer technique consisting of three steps: mouse ventral laparotomy, transuterine microinjection, and in vivo electroporation. We use digital video microscopy to demonstrate the critical experimental embryological techniques.

The mammalian inner ear has 6 distinct sensory epithelia: 3 cristae in the ampullae of the semicircular canals; maculae in the utricle and saccule; and ...

Patch Clamp Recordings from Embryonic Zebrafish Mauthner Cells

Mauthner cells (M-cells) are large reticulospinal neurons located in the hindbrain of teleost fish. They are key neurons involved in a characteristic behavior known as the C-start or escape response that occurs when the organism perceives a threat. The M-cell has been extensively studied in adult goldfish where it has been shown to receive a wide range of excitatory, inhibitory and neuromodulatory signals1. We have been examining M-cell activity in embryonic zebrafish in order to study aspects of synaptic development in a vertebrate preparation. In the late 1990s Ali and colleagues developed a preparation for patch clamp recording from M-cells in zebrafish embryos, in which the CNS was largely intact2,3,4. The objective at that time was to record synaptic activity from hindbrain neurons, spinal cord neurons and trunk skeletal muscle while maintaining functional synaptic connections within an intact brain-spinal cord preparation. This preparation is still used in our laboratory today. To examine the mechanisms underlying developmental synaptic plasticity, we record excitatory (AMPA and NMDA-mediated)5,6 and inhibitory (GABA and glycine) synaptic currents from developing M-cells. Importantly, this unique preparation allows us to return to the same cell (M-cell) from preparation to preparation to carefully examine synaptic plasticity and neuro-development in an embryonic organism. The benefits provided by this preparation include 1) intact, functional synaptic connections onto the M-cell, 2) relatively inexpensive preparations, 3) a large supply of readily available embryos 4) the ability to return to the same cell type (i.e. M-cell) in every preparation, so that synaptic development at the level of an individual cell can be examined from fish to fish, and 5) imaging of whole preparations due to the transparent nature of the embryos.

We have developed an intact brain-spinal cord preparation to record and monitor electrical activity via patch clamp recording from the Mauthner neurons and other reticulospinal cells in zebrafish embryos. Thus, we are able to record excitatory and inhibitory synaptic currents, voltage-gated channel activity and action potentials from key neurons in a developing embryo.

Mauthner cells (M-cells) are large  reticulospinal neurons located in the hindbrain of teleost fish. They are key  neurons involved in a characteristic ...

Whole-cell Patch-clamp Recordings from Morphologically- and Neurochemically-identified Hippocampal Interneurons

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal population (10-20%), but show a high level of physiological, morphological, and neurochemical heterogeneity, reflecting their diverse functions. Therefore, investigation of interneurons provides important insights into the organization principles and function of neuronal circuits. This, however, requires an integrated physiological and neuroanatomical approach for the selection and identification of individual interneuron types. Whole-cell patch-clamp recording from acute brain slices of transgenic animals, expressing fluorescent proteins under the promoters of interneuron-specific markers, provides an efficient method to target and electrophysiologically characterize intrinsic and synaptic properties of specific interneuron types. Combined with intracellular dye labeling, this approach can be extended with post-hoc morphological and immunocytochemical analysis, enabling systematic identification of recorded neurons. These methods can be tailored to suit a broad range of scientific questions regarding functional properties of diverse types of cortical neurons.

Cortical networks are controlled by a small, but diverse set of inhibitory interneurons. Functional investigation of interneurons therefore requires targeted recording and rigorous identification. Described here is a combined approach involving whole-cell recordings from single or synaptically-coupled pairs of neurons with intracellular labeling, post-hoc morphological and immunocytochemical analysis.

GABAergic inhibitory interneurons play a central role within neuronal circuits of the brain. Interneurons comprise a small subset of the neuronal ...

Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with simultaneous intracellular biocytin filling allows a correlated analysis of their structural and functional properties. With this method it is possible to identify and characterize both pre- and postsynaptic neurons by their morphology and electrophysiological response pattern. Paired recordings allow studying the connectivity patterns between these neurons as well as the properties of both chemical and electrical synaptic transmission. Here, we give a step-by-step description of the procedures required to obtain reliable paired recordings together with an optimal recovery of the neuron morphology. We will describe how pairs of neurons connected via chemical synapses or gap junctions are identified in brain slice preparations. We will outline how neurons are reconstructed to obtain their 3D morphology of the dendritic and axonal domain and how synaptic contacts are identified and localized. We will also discuss the caveats and limitations of the paired recording technique, in particular those associated with dendritic and axonal truncations during the preparation of brain slices because these strongly affect connectivity estimates. However, because of the versatility of the paired recording approach it will remain a valuable tool in characterizing different aspects of synaptic transmission at identified neuronal microcircuits in the brain.

Patch-clamp recordings and simultaneous intracellular biocytin filling of synaptically coupled neurons in acute brain slices allow a correlated analysis of their structural and functional properties. The aim of this protocol is to describe the essential technical steps of electrophysiological recording from neuronal microcircuits and their subsequent morphological analysis.

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding these fundamental functions will shed light on the information transfer in these neurons. Dendritic patch-clamp recordings provide the possibility to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels. However, these fine structures are not easily accessible to patch pipettes because of their small diameter. This report describes a step-by-step procedure to collect stable and reliable recordings from the dendrites of dopaminergic neurons in acute slices. Electrophysiological measurements are combined with post hoc recovery of cell morphology. Successful experiments rely on improved preparation of slices, solutions and pipettes, adequate adjustment of the optics and stability of the pipette in contact with the recorded structure. Standard principles of somatic patch-clamp recording are applied to dendrites but with a gentler approach of the pipette. These versatile techniques can be implemented to address various questions concerning the excitable properties of dendrites.

Subcellular patch-clamp recordings offer the possibility to investigate the functional properties of dendrites. However these fine structures are not easily accessible due to their small diameter. The protocol described here aims to facilitate the collection of stable and reliable recordings from the dendrites of dopamine neurons in slices in vitro.

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding ...